NADP(H)-dependent biocatalysis without adding NADP(H)

Significance Within cells, enzymes and cofactors catalyzing multistep processes (cascades) are often confined together—either in enclosures (e.g., organelles) or via physical association (metabolons). Nanoconfinement, offering potential general advantages for catalysis, can also be achieved by loading enzymes and their exchangeable cofactors into a porous, electrically conducting inorganic material, thereby enabling catalysis to be channeled, energized, and investigated electrochemically. Such nanoconfinement enables a cascade comprising electroactive ferredoxin NADP+ reductase and isocitrate dehydrogenase to be active for days, catalyzing exhaustive oxidation of bulk isocitrate by recycling trapped NADP(H) carried in on isocitrate dehydrogenase. Nanoconfinement massively increases the efficiency of cofactor-dependent cascade catalysis and has conceptual relevance for prebiotic evolution where complex organic molecules might have formed in gaps and cracks of minerals.


Electrode Fabrication
Nanoporous indium tin oxide (ITO) electrodes were made by electrophoretic deposition of ITO nanoparticles (<50 nm, Sigma-Aldrich) onto either pyrolytic graphite edge (PGE) or titanium foil electrodes as previously described (1), except that the deposition time was increased to 8-12 minutes from 6 minutes in order to produce a thicker ITO layer.

Enzyme Loading
Enzymes were loaded onto nanoporous ITO electrodes as previously described (1). Briefly, a concentrated mixed enzyme solution (4 µL for 0.03 or 0.06 cm 2 PGE electrodes; 18 µL for 4 cm 2 Ti foil electrode) was incubated on the electrode for ~45 minutes at room temperature before being rinsed thoroughly to ensure that any unbound enzyme was removed. For PGE electrodes, 0.85 nmoles of IDH1 (both wild-type and R132H) was used, and the amount of FNR was adjusted to achieve the desired molar loading ratio. Four-fold more enzyme (3.4 nmoles wild-type IDH1 and 6.8 nmoles FNR) was used to load the scaled-up 4 cm 2 Ti foil electrode described in Figure 3. Enzyme molar ratios were calculated based on IDH1 homodimers.

Electrochemical Quantification of Adsorbed FNR
Electroactive FNR was quantified using cyclic voltammetry as previously described (1).

Electrochemical Experiments
Electrochemical experiments were performed in an anaerobic glove box (Braun Technologies) with a nitrogen atmosphere (O2 < 1 ppm) using an Autolab PGSTAT 10 potentiostat and Nova software. In-house custom glass cells were used for all experiments; a two-chamber glass cell was used for experiments with an ITO/PGE rotating disc electrode (the working electrode and platinum counter electrode were in the same chamber) (1, 3), while a three-chamber glass cell was used for the scaled-up experiment with the working, counter, and reference electrodes housed in separate chambers (4). In the scaled-up experiment (threechamber cell), the counter electrode and working electrode chambers were separated by a glass frit and contained the same buffer solution. In all experiments, the reference electrode chamber contained 0.1 M NaCl and was connected to the working electrode chamber via a Luggin capillary. Electrode potentials were measured against a saturated calomel electrode (SCE) and converted to SHE using a SCE to SHE conversion table (1).

Live Buffer Exchange Protocol
Live buffer exchanges (i.e., where current is continuously monitored during the buffer exchange) were performed as previously described (1), except that different volumes of exchange buffer were used depending on how much dilution was required. Briefly, a 1 mL syringe was used to reduce the reaction volume to ~1.5 mL while making sure that the electrode was always submerged in order to maintain electrical contact. A 60 mL syringe containing fresh buffer solution (30 mL buffer for >1000-fold dilution or 47 mL for ~55000-fold dilution) was then used to slowly inject 8 mL of buffer into the cell. The syringe was removed, and a 20 mL syringe was then used to remove 8 mL of buffer from the cell now containing the diluted starting buffer. Once the buffer volume was reduced to ~1.5 mL again, 8 mL of buffer solution was again added to the cell, and the process was repeated until all of the exchange buffer was used and the final cell volume was equal to the starting volume. The electrode rotation (1000 rpm) provided adequate solution agitation to ensure mixing during the procedure.

IDH1-FNR Experiments in Dilute Solution
The isocitrate oxidation activity of the IDH1-FNR cascade was measured in dilute solution both with added NADPH (positive control) and without (only IDH1-copurified NADP(H) present) over 12 hours for comparison with the same cascade reaction nanoconfined in electrodes. Two experiments were carried out at different enzyme concentrations based on the amount of FNR and IDH1dimer that was quantified on individual small (0.06 cm 2 ) and large (4 cm 2 ) electrodes used in experiments (see Supporting Figure 11 for details), equivalent to 2.6 (2.7) and 40 (41.5) nM of FNR (parentheses denote [IDH1dimer]) in a 4 mL solution. The solution reactions were allowed to run at 25 °C for 12 h before heat quenching and analysis by 1 H NMR. NADP + was recycled by FNR for use by IDH1, with a large excess of benzyl viologen in solution (25 mM) to continuously re-oxidize FNR in place of the electrode. The cascade activity in solution was determined based on the amount of product made (2OG) in the solution over a 12-hour period (quantified by 1 H NMR after heat quenching). The molar ratio of FNR:IDH1dimer used was 1:1.04 ([IDH1] = 1.04 x [FNR]), the same ratio that was quantified in the electrode nanopores for the relevant electrochemical experiments using the FNR:IDH1dimer = 2:1 loading ratio ( Table 1). Other conditions: 400 µL reaction volume, 100 mM HEPES (pH 8), 10 mM MgCl2, 5 mM D-isocitrate, 25 mM benzyl viologen, 25 °C. All reactions were performed in triplicate with error bars representing the standard deviation.

H Nuclear Magnetic Resonance (NMR) experiments
NMR spectra were obtained using a Bruker AVIII 700 MHz NMR spectrometer equipped with an inverse 5mm TCI 1 H/ 13 C/ 15 N cryoprobe. The water signal was suppressed by excitation sculpting. For copurification experiments, proteins were buffer exchanged into NMR buffer (50 mM Tris-d11, 100 mM NaCl, 10 % D2O, pH 7.5) using Micro Bio-Spin 6 columns (Bio-Rad) according to the manufacturer's protocol. The final protein concentrations were 50 µM for FNR and IDH1 R132H and 124 µM for wild-type IDH1 (a higher concentration was required to compensate for the weaker NADP(H) signal intensity due to copurification with both NADP + and NADPH). A 1 H spectrum (NS: 64, relaxation delay: 2 s) of this solution was obtained before the protein was denatured by heat (2 min, 100 °C). A spectrum of the denatured protein sample was recorded and the copurifying molecules identified by spiking with 100 µM of NADPH, NADP + , or FAD.
A control experiment to investigate oxidation of NADPH to NADP + in the presence of FAD was set up according to the same parameters. The sample was composed of FAD (1 mM) and NADPH (1 mM) in NMR buffer. An equivalent experiment was set up with benzyl viologen (1 mM) and NADPH (1 mM).
For quantitative NMR experiments, 80 µL of a sample from electrochemical experiments in 100 mM HEPES was mixed with 16 µL D2O (final 10% concentration), 16 µL 3-(Trimethylsilyl)propionic-2,2,3,3-d4 acid sodium salt solution (TSP; final: 1 mM), 48 µL MQ water. The concentration of D-isocitrate and 2OG was calculated using the concentration of the internal standard TSP. The number of scans was 32, and D1 was 30 s. If the sample concentration was small, the number of scans was increased to 128. The water signal was suppressed by excitation sculpting.

Non-denaturing Mass Spectrometry Experiments
Non-denaturing mass spectrometry studies were conducted as previously reported (2). In brief, the protein samples were exchanged into buffer (ammonium acetate, 200 mM, pH 7.5) with Micro Bio-Spin 6 Columns (Bio-Rad). The samples were analysed at a concentration of 20 µM on a quadrupole-TOF (Waters Synapt G2Si) instrument and an Advion Triversa Nanomate chip-based ESI autosampler. The parameters were as follows: 1.7-1.8 kV, spray backing gas pressure 0.6 psi, inlet pressure 3.7 mbar; cone voltages 100V and 5.2 V.

Derivation of Equations 1 and 2 (main text)
In order to derive an equation giving the current for an electroactive enzyme as a function of substrate concentration, we start with the equation describing the current from an electroactive enzyme, = Γ T , Supporting Eq. 1 where I is the current (Amps), n is the number of electrons transferred per molecule, A is the electrode surface area (cm 2 ), F is the Faraday constant (96,485 C per mole of electrons), Γ is the enzyme coverage (moles cm -2 ), and kT is the enzyme turnover frequency (s -1 ). Because the enzyme turnover frequency, kT, changes with substrate concentration as described by the  To separate the overlapping peaks in the bottom panel (wild-type IDH1) and allow integration, the exponential parts of each peak was fit using an asymptotic exponential equation (approaching 0). The decay rate constant used to fit the first two peaks (red and blue) was determined by fitting the same exponential equation to the green peak (2 molecules of bound NADP(H)), the trailing tail of which does not overlap with other peaks.

Supporting
Supporting Figure 3. 1 H NMR data showing that no NADP(H) is copurified with FNR. The FAD cofactor only becomes visible after the enzyme is denatured, which was confirmed by adding additional FAD (100 µM) to the NMR tube containing the denatured FNR. Interestingly, the NADPH that was added to the sample (100 µM) was oxidized to NADP + despite the FNR being denatured. A control experiment (Supporting Figure 4) confirmed that "free" FAD in solution is able to directly oxidize NADPH. , it was observed that the fully-oxidized FAD peak (denoted with a yellow star) was much smaller and broader than expected, likely due to it having been reduced by NADPH. After 12 hours, the oxidized FAD peak can be clearly seen, having been reoxidized by oxygen, alongside NADP + (no NADPH remains). (B) Benzyl viologen is not able to oxide NADPH in solution. The bottom scan is after 15 minutes of incubation; the top scan is after 12 hours.

Supporting
Supporting Figure 5. Bar chart showing no significant difference (p = 0.45; two-tailed t-test assuming unequal variances) in the amount of enzyme rinsed from 0.06 cm 2 electrodes (3) after enzyme loading vs control samples (3), where no electrode was used (directly diluted enzyme stock). The three electrodes were loaded with enzyme (1.27 nmoles total; FNR:IDH1 R132Hdimer; 1:2) following the typical procedure (see Materials and Methods).
The electrodes were rinsed using buffer (100 mM HEPES (pH = 8)), and the rinsed solution was kept for comparison with three aliquots of the same amount of enzyme that was directly diluted into the same amount of buffer used for rinsing. Based on absorbance at 280 nm, the rinsed solution contains roughly the same amount of enzyme as the directly diluted stocks, confirming that only a tiny fraction of the enzyme used is actually taken up into the electrode nanopores with the vast majority being rinsed off. Each sample was analyzed using three technical replicates. Error bars represent the standard deviation.